Process of preparing 3d array of particles and exemplary application thereof in sensor fabrication

ABSTRACT

The present invention provides a novel and efficient process of preparing a highly organized 3D array of particles by stacking multiple 2D arrays of the particles. The 3D array of particles so prepared is used in fabrication of sensors, such as molecular imprinted photonic (MIP) crystal sensor. The sensor has a 3D array of voids each having a void internal wall. The void internal walls have cavities each having a cavity internal wall made from a material containing the non-metallic element. A binding of the analytes to the cavities induces a detectable variation of the optical property of the 3D array of voids. The invention exhibits numerous technical merits such as high sensitivity, high specificity, fast detection, ease of operation, low power consumption, zero chemical release, and low operation cost, among others.

CROSS-REFERENCE TO RELATED APPLICATIONS

The present application for patent claims the benefit of U.S. Provisional Patent Application No. 62/973,591 filed Oct. 15, 2019, the entire disclosures of which is incorporated herein by reference.

STATEMENT REGARDING FEDERALLY SPONSORED RESEARCH OR DEVELOPMENT

The invention was made with the US EPA Small Business Innovation Research (SBIR) support under Contract No. 68HERD19C0010. The government has certain rights in the invention.

FIELD OF THE INVENTION

The present invention generally relates to a general process of preparing a 3D array of particles. Although the invention will be illustrated, explained and exemplified by a 3D array of silica nanoparticles, it should be appreciated that the present invention can also be applied to other fields, for example, polystyrene nanoparticles, polymer beads, oxides, oxide ceramic nanoparticles, nitrides, carbides, quantum dots, macromolecules, carbon nanostructures, and the like. Although the usefulness of the invention is found in an exemplary field of sensor fabrication, such as molecular imprinted photonic (MIP) crystal sensor fabrication, it should be appreciated that the usefulness of the present invention can also be found in many other micro-fabrication fields.

BACKGROUND OF THE INVENTION

There are many kinds of 3D array of entities. For example, the constituents such as atoms, molecules, or ions in a crystal or crystalline solid are arranged in a highly ordered microscopic structure, forming a crystal lattice that extends in all directions. Crystals can be prepared from precipitating from a solution, freezing, or deposition directly from a gas. Crystallization starts from nucleation characterized by the appearance of a crystalline phase from either a super cooled liquid or a supersaturated solvent. Then, crystal growth will increase the size of particles and lead to a crystal state.

Examples of self-assembly in material science include the formation of molecular crystals, colloids, lipid bilayers, phase-separated polymers, and self-assembled monolayers. Molecular self-assembly is the process by which molecules adopt a defined arrangement without guidance or management from an outside source. Self-assembled monolayers (SAM) of organic molecules are molecular assemblies formed spontaneously on surfaces by adsorption and are organized into more or less large ordered domains.

However, there remains a need for novel methods of fabricating 3D array of silica nanoparticles, polystyrene nanoparticles, polymer beads, oxides, oxide ceramic nanoparticles, nitrides, carbides, quantum dots, macromolecules, carbon nanostructures, and the like.

On the other hand, there also exists a need for sensors and sensing devices used for detecting or measuring analytes containing a non-metallic element. For example, compounds from a large family of perfluorinated chemicals (PFCs), such as perfluorooctane sulphonate (PFOS) and perfluorooctanoic acid (PFOA), have attracted worldwide attention in the scientific regulatory community and among the public due to their persistent, bio-accumulative, and toxic characteristics that can significantly deteriorate human health. PFOS and PFOA have found significant usage in many industrial and consumer applications that require high chemical stability and dirt-water-oil repellency, characteristics which are provided by the strong electro-negativity and small atomic size of fluorine molecules. They are also used for firefighting at airfields because of their inherent ability to create aqueous firefighting form foams (AFFFs) to extinguish fuel and hydrocarbon fires. Unfortunately, the chemical nature of fluorine makes the carbon-fluorine bond the strongest in nature, which makes these fluorinated compounds resistant to chemical or biochemical reactions and degradation processes. Due to increasing concerns over the long-term health effects of PFOS and PFAS on the human body, regulatory agencies have set limits for the concentrations of PFOS and PFAS in drinking water. In 2016, the United States Environmental Protection Agency (USEPA) established a lifetime health advisory (LHA) level of 70 parts per trillion (ppt) for individual or combined concentrations of PFOA and PFOS in drinking water. Recent studies indicate that exposure to PFOA and PFOS over certain levels may result in adverse health effects, including developmental defects in fetuses and breastfed infants, cancer, liver effects, immune effects, thyroid effects, and others. Hence, the development of trace detection and monitoring systems for PFOS and PFOA in water is highly necessary.

Currently, mass-spectrometry-based technologies are the main methods used to detect trace perfluorinated acids in various samples with sufficient sensitivity and selectivity. However, these methods require large and expensive equipment, have high operation costs, and sometimes suffer matrix interferences, making them unsuitable for routine analysis of PFOS and PFOA in the field.

Lab analysis for PFAS (EPA 537) is time-consuming and expensive, taking as long as 3 weeks and costing up to $450 per sample. Mobile labs can be rented for ˜$500/week to cut down on analysis time. The detection of PFAS compounds in the field remains a big problem to solve. People currently send all samples back to a lab, which is time-consuming and expensive and creates bottlenecks for fairly large projects.

Advantageously, the present invention provides a novel method of fabricating 3D array of silica nanoparticles, polystyrene nanoparticles, polymer beads, oxides, oxide ceramic nanoparticles, nitrides, carbides, quantum dots, macromolecules, carbon nanostructures, and the like, but also a novel sensor and a sensing device that exhibit numerous technical merits. For example, the sensor and sensing device of the invention is fieldable, fast (minutes vs two weeks for lab measurement), and much cheaper ($20 to $30 per sample vs $200 to $300 per sample for lab measurement) than the techniques currently on the market.

SUMMARY OF THE INVENTION

One aspect of the present invention provides a process of preparing a 3D array of particles by stacking multiple 2D arrays of the particles. The process may include:

(a) providing a container with a substrate inside the container;

(b) introducing a first liquid into the container to immerse the substrate, and optionally introducing a second liquid to float on top surface of the first liquid and form a two-phase system;

(c) assembling the particles on top surface of the first liquid to form a monolayer (2D array) of the particles thereon, or optionally assembling the particles to form a monolayer at an interface between the first liquid and the second liquid;

(d) moving the monolayer and/or the substrate to reduce a distance therebetween until the monolayer is deposited on the substrate;

(e) subjecting the substrate with the monolayer deposited thereon to steps (a)-(d) to stack/deposit another monolayer on top of the monolayer previously deposited; and

(f) optionally repeating step (e) until a desired number of monolayers are stacked on the substrate.

In representative embodiments, the process is implemented with the following steps:

(i) providing a container with a substrate on a bottom of the container;

(ii) filling a first liquid into the container until the substrate is immersed in the first liquid;

(iii) adding a preparation of the particles into the container, and assembling the particles on top surface of the first liquid to form a monolayer (2D array) of the particles thereon;

(iv) removing or discharging the first liquid from the container so that said monolayer of the particles falls down onto the substrate, and is deposited thereon;

(v) refilling the first liquid into the container until the substrate and the particles previously deposited thereon is immersed in the first liquid;

(vi) repeating steps (iii) and (iv) so that another monolayer of the particles is deposited on the substrate by stacking over an immediate monolayer that has previously deposited thereon; and

(vii) optionally repeating steps (v) and (vi) until a desired number of monolayers of the particles are deposited on the substrate.

Another aspect of the invention provides a process for preparing a sensing body of a working sensor for a sensing device useful for detecting an analyte containing a non-metallic element. The process includes:

(1) fabricating a 3D array of particles as described above;

(2) infiltrating the 3D array of particles with a mixture of a solidifiable material and the analyte, wherein the solidifiable material comprises said non-metallic element too;

(3) solidifying the solidifiable material with the analyte; and

(4) washing away the 3D array of particles and the analyte, forming a sensing body including a 3D array of voids each having a void internal wall.

At least a part of the voids so formed are interconnected to each other and are configured to expose to said analyte in a future sample, and admit said analyte into said at least a part of the voids. Void internal walls of said at least a part of the voids have cavities each having a cavity internal wall. Each of the cavities has a shape that is complementary to a shape of the analyte. The cavity internal wall is made from a material containing said non-metallic element.

Still another aspect of the invention provides a sensing device comprising a working sensor for detecting an analyte containing a non-metallic element. The working sensor comprises a sensing body including a 3D array of voids each having a void internal wall. At least a part of the voids are interconnected to each other and are configured to expose to the analyte, and admit the analyte into the at least a part of the voids. Void internal walls of the at least a part of the voids have cavities each having a cavity internal wall. Each of the cavities has a shape that is complementary to a shape of the analyte. The cavity internal wall is made from a material containing the non-metallic element.

A further aspect of the invention provides a method of measuring an analyte containing a non-metallic element using the aforementioned sensing device. The method includes (i) contacting a sample of the analyte with the working sensor, (ii) binding the analyte to the cavities and inducing or triggering a detectable variation of the optical property of the 3D array of voids, including a spectrum of light that is transmitted through, reflected from, and/or diffracted from the 3D array of voids, and (iii) correlating a degree of the detectable variation to an amount of the analytes bound to the cavities.

The above features and advantages and other features and advantages of the present invention are readily apparent from the following detailed description of the best modes for carrying out the invention when taken in connection with the accompanying drawings.

BRIEF DESCRIPTION OF THE SEVERAL VIEWS OF THE DRAWINGS

The present invention is illustrated by way of example, and not by way of limitation, in the figures of the accompanying drawings and in which like reference numerals refer to similar elements. All the figures are schematic and generally only show parts which are necessary in order to elucidate the invention. For simplicity and clarity of illustration, elements shown in the figures and discussed below have not necessarily been drawn to scale. Well-known structures and devices are shown in simplified form, omitted, or merely suggested, in order to avoid unnecessarily obscuring the present invention.

FIG. 1A is a flow chart showing a general process of preparing a 3D array of particles in accordance with an exemplary embodiment of the present invention.

FIG. 1B schematically illustrates the general process of preparing a 3D array of particles in accordance with an exemplary embodiment of the present invention.

FIG. 1C is a flow chart showing a specific process of preparing a 3D array of particles in accordance with an exemplary embodiment of the present invention.

FIG. 1D schematically illustrates the specific process of preparing a 3D array of particles in accordance with an exemplary embodiment of the present invention.

FIG. 1E is a flow chart showing another specific process of preparing a 3D array of particles in accordance with an exemplary embodiment of the present invention.

FIG. 1F schematically illustrates the another specific process of preparing a 3D array of particles in accordance with an exemplary embodiment of the present invention.

FIG. 1G illustrates a step of compressing a monolayer of particles with barriers in accordance with an exemplary embodiment of the present invention.

FIG. 1H is a flow chart showing a process for preparing a sensing body in accordance with an exemplary embodiment of the present invention.

FIG. 1I schematically illustrates a process for preparing a sensing body in a working sensor in accordance with an exemplary embodiment of the present invention.

FIG. 1J schematically shows a sensing device comprising the working sensor in accordance with an exemplary embodiment of the present invention.

FIG. 2A illustrates a sensing device comprising a working sensor and a reference sensor in accordance with an exemplary embodiment of the present invention.

FIG. 2B is a flow chart of a method for measuring an analyte containing a non-metallic element in accordance with an exemplary embodiment of the present invention.

FIG. 3 illustrates a photonic crystal sensing chip in accordance with an exemplary embodiment of the present invention.

FIG. 4 demonstrates the fabrication procedure for a hierarchical porous sensor chip sensing chip in accordance with an exemplary embodiment of the present invention.

FIG. 5 depicts a colloid crystal template prepared using water-hexane interfacial assembly in accordance with an exemplary embodiment of the present invention.

FIG. 6 shows images and size distribution of silica nanoparticle colloids in accordance with an exemplary embodiment of the present invention.

FIG. 7 shows SEM surface morphologies of 1-layer and 10-layer samples in accordance with an exemplary embodiment of the present invention.

FIG. 8 shows the UV-Vis absorption spectra of a 5-layer and a 10-layer silica colloid crystal template with a particle size of 180 nm in accordance with an exemplary embodiment of the present invention.

FIG. 9 shows vis-NIR spectra of a colloid crystal sample measured after assembly of 5 layers and 10 layers in accordance with an exemplary embodiment of the present invention.

FIG. 10 illustrates a fabrication process for PFOA-imprinted inverse polymer opal structure in accordance with an exemplary embodiment of the present invention.

FIG. 1I shows a fabricated inverse opal sensor from a colloid crystal template of 300 nm nanoparticles in accordance with an exemplary embodiment of the present invention.

FIG. 12 shows FTIR spectra of a UV-cured polymer (red) and a mixed precursor solution (black) in accordance with an exemplary embodiment of the present invention.

FIG. 13 shows surface morphologies of the fabricated reverse opal polymer structures at different magnifications in accordance with an exemplary embodiment of the present invention.

FIG. 14 shows vis-NIR spectra of the fabricated inverse opal sensor after HF etching (red) and after PFOA removal (green) in accordance with an exemplary embodiment of the present invention.

FIG. 15A shows vis-NIR spectra of a sensor at analyte concentrations of from 10 ppt to 10,000 ppt in accordance with an exemplary embodiment of the present invention.

FIG. 15B shows a calibration curve in the range from 0 ppt to 10,000 ppt in accordance with an exemplary embodiment of the present invention.

FIG. 16 shows vis-NIR spectra of non-molecularly imprinted sensors at PFOA concentrations of from 10 ppt to 1,000 ppt in accordance with an exemplary embodiment of the present invention.

FIG. 17 shows a chip assembled onto a clear microscope slide support in accordance with an exemplary embodiment of the present invention.

FIG. 18 shows the UV-Vis spectra of colloid crystals independently (Green and Red) and when they are combined (yellow) in accordance with an exemplary embodiment of the present invention.

FIG. 19 shows Vis-NIR spectra from 4 corner areas of a larger 3D array of particles in accordance with an exemplary embodiment of the present invention.

DETAILED DESCRIPTION OF THE PREFERRED EMBODIMENT

In the following description, for the purposes of explanation, numerous specific details are set forth in order to provide a thorough understanding of the present invention. It is apparent, however, to one skilled in the art that the present invention may be practiced without these specific details or with an equivalent arrangement.

Where a numerical range is disclosed herein, unless otherwise specified, such range is continuous, inclusive of both the minimum and maximum values of the range as well as every value between such minimum and maximum values. Still further, where a range refers to integers, only the integers from the minimum value to and including the maximum value of such range are included. In addition, where multiple ranges are provided to describe a feature or characteristic, such ranges can be combined.

It is also to be understood that the terminology used herein is for the purpose of describing particular embodiments only, and is not intended to limit the scope of the invention. For example, when an element is referred to as being “on”, “connected to”, or “coupled to” another element, it can be directly on, connected or coupled to the other element or intervening elements may be present. In contrast, when an element is referred to as being “directly on”, “directly connected to”, or “directly coupled to” another element, there are no intervening elements present.

FIG. 1A is a flow chart showing a general process of preparing the 3D array of particles by stacking multiple 2D arrays of the particles. The general process includes step (a) providing a container 61 with a substrate 62 inside the container 61, as shown in FIG. 1B. Step (b) is introducing a first liquid 63 into the container to immerse the substrate 62, and optionally introducing a second liquid 68 to float on top surface of the first liquid 63 and form a two-phase system. Step (c) is assembling the particles on top surface of the first liquid 63 to form a monolayer 65 (2D array) of the particles thereon, or optionally assembling the particles to form a monolayer 65 at an interface between the first liquid 63 and the second liquid 68. Step (d) is reducing a distance between the monolayer 65 and the substrate 62, by relatively moving them, for example, lowering the monolayer 65 only as shown in Panel (A) of FIG. 1B, lifting up the substrate 62 only as shown in Panel (B) of FIG. 1B, or moving both the monolayer 65 and the substrate 62 toward each other as shown in Panel (C) of FIG. 1B, until the monolayer is deposited on the substrate. In step (e), the substrate with the monolayer deposited thereon is subjected to steps (a)-(d) so as to stack/deposit another monolayer on top of the monolayer 65 previously deposited. Optional step (f) is repeating step (e) until a desired number of monolayers are stacked on the substrate 62, for example repeating step (e) for 1 to 1000 times, 2 to 500 times, 5 to 100 times, or 10 to 50 times. A flat surface (e.g. upper surface) of the substrate 62 for depositing the monolayer(s) is not vertical to the top surface of the first liquid. For example, the angle between the two surfaces may range from −89° to 89°, from −70° to 70°, from −50° to 50°, from −35° to 35°, from −20° to 20°, from −10° to 10°, from −5° to 5° such as substantially 0°. In the following, reprehensive embodiments with the angle of substantially 0° will be described in more details.

FIG. 1C is a flow chart showing a specific process of preparing the 3D array of particles. The specific process includes step (i) providing a container 61 with a substrate 62 on a bottom of the container 61, as shown in FIG. 1D. Step (ii) is filling a first liquid 63 into the container 61 until the substrate 62 is immersed in the first liquid 63. Step (iii) is adding a preparation of the particles 64 into the container 61, and assembling the particles 64 on top surface of the first liquid 63 to form a monolayer 65 (2D array) of the particles thereon. The monolayer may be formed under an activation of vibration energy, which can be accomplished by mechanical vibration and/or ultrasonic vibration. Step (iv) is removing or discharging the first liquid 63 from the container 61 so that said monolayer of the particles 65 falls down onto the substrate 62, and is deposited substrate 62. An optional heating step may be employed to strengthen the binding between the monolayer of the particles 65 and the substrate 62. Step (v) is refilling the first liquid 63 into the container 61 until the substrate 62 and the particles previously 65 deposited thereon is immersed in the first liquid 63. Step (vi) is repeating steps (iii) and (iv) so that another monolayer of the particles 66 is deposited on the substrate 62 by stacking over an immediate monolayer 65 that has previously deposited thereon. Optional step (vii) is repeating steps (v) and (vi) until a desired number of monolayers (65, 66, 67 . . . ) of the particles are deposited on the substrate 62.

FIG. 1E shows a specific embodiment of the process of FIG. 1C for preparing the 3D array of particles by stacking multiple 2D arrays of the particles. As shown in FIG. 1E and FIG. 1F, step (i) providing the container 61 with the substrate 62 on the bottom of the container. Step (ii) is filling the first liquid 63 into the container 61 until the substrate 62 is immersed in the first liquid 63. An additional step (ii.5) is adding a second liquid 68 into the container 61, so that the second liquid 68 is floating on top of the first liquid 63. An interface is formed between the two liquids (63, 68). Step (iii) is adding the preparation of the particles 64 into the container 61, and assembling the particles 64 between the first liquid 63 and the second liquid 68 (or at their interface) to form the monolayer 65 (2D array) of the particles. The monolayer may be formed under an activation of vibration energy, which can be accomplished by mechanical vibration and/or ultrasonic vibration. Step (iv) is removing or discharging both the first liquid 63 and the second liquid 68 from the container 61 so that said monolayer 65 of the particles falls down onto the substrate 62, and is deposited thereon 62. Step (v) is refilling the first liquid 63 into the container 61 until the substrate 62 and the particles 65 previously deposited thereon is immersed in the first liquid 63. Step (vi) is repeating steps (ii.5), (iii) and (iv) so that another monolayer (not shown) of the particles is deposited on the substrate 62 by stacking over the immediate monolayer 65 that has previously deposited thereon. An optional step (vii) is repeating steps (v) and (vi) until a desired number of monolayers of the particles are deposited or stacked on the substrate 62, similar to that shown in FIG. 1D.

In various embodiments of the invention, the substrate 62 comprises a material made of glass or oxide ceramic. The first liquid comprises a hydrophilic liquid such as water, and the second liquid comprises a hydrophobic liquid or oil such as hydrocarbon liquid such as hexane, heptane, or any mixture thereof. The particles 64 may comprise silica nanoparticles, polystyrene nanoparticles, polymer beads, oxides, oxide ceramic nanoparticles, nitrides, carbides, quantum dots, macromolecules, carbon nanostructures, or any mixture thereof. The particles assembled in one of the monolayers (2D arrays) may be the same as, or different from, the particles assembled in another one of the monolayers (2D arrays). Moreover, some particles assembled in a monolayer may be the same as, or different from, other particles assembled in the same monolayers. The preparation of the particles 64 may be a suspension of the particles in an organic solvent such as ethanol, for example nanoparticle colloid mono-dispersed in ethanol.

The particles 64 may have an average size in the range of 10-1000 nm, 50-1000 nm, 50-500 nm, 150-300 nm, or 180-400 nm. Each monolayer of the particles has a thickness that is substantially the same as the particles' average size. The finished stack of the monolayers may include from 5 to 20, e.g. 10 monolayers, and the stack may have a height of approximately 2-10 μm.

Referring back to FIG. 1E, Step (iii) may further include an additional step, for example, compressing the monolayer of the particles with a pair of barriers 78 to reduce the area of the monolayer and to pack the particles 64 in the monolayer more densely or more intimately, as shown in FIG. 1G. Moreover, adding the preparation of the particles into the container in step (iii) may be accomplished with a pipette, a syringe pump, or any combination thereof. The entire process as shown in FIG. 1E may further include a step of (viii), annealing the deposited particles at an elevated temperature such as 50-100° C. e.g. 70° C. for a period of time such as 5-20 minutes e.g. 10 minutes, to evaporate a residue of the first liquid, the second liquid, and a liquid in the preparation of the particles, optionally wherein the annealing is accomplished by oven drying, IR lamp heating, or any combination thereof.

FIG. 1H is a flow chart showing a process for preparing a sensing body of a working sensor for a sensing device that is useful for detecting an analyte containing a non-metallic element. As shown in FIG. 1H and FIG. 1I, Step (1) is fabricating a 3D array of particles 71 as described above on substrate 62. Step (2) is infiltrating the 3D array of particles 71 with a mixture 72 of a solidifiable material and the analyte. The solidifiable material comprises said non-metallic element too. An optional base 73 may be used to seal and support the side of the 3D array 71 that is opposite to the side covered by substrate 62. Step (3) is solidifying the solidifiable material with the analyte. Step (4) washing away the 3D array of particles 71 and the analyte, forming a sensing body including a 3D array 74 of voids 75 each having a void internal wall 76. At least a part of the voids are interconnected to each other and are configured to expose to the analyte in a future sample, and to admit the analyte into the at least a part of the voids. Void internal walls 76 of said at least a part of the voids 75 have cavities 77 each having a cavity internal wall. Each of the cavities 77 has a shape that is complementary to a shape of the analyte. In various embodiments, the cavity internal wall is made from a material containing the non-metallic element.

With reference to FIG. 1J, various embodiments of the invention provide a sensing device 001 comprising a working sensor 100 for detecting an analyte 10 containing a non-metallic element. The sensor 100 comprises a sensing body 110 including a 3D array of voids 120 each having a void internal wall 121. At least a part of the voids 120 are interconnected to each other and are configured to expose to the analyte 10, and admit the analyte 10 into the at least a part of the voids 120. Void internal walls 121 of the at least a part of the voids 120 have cavities 130 each having a cavity internal wall 131. Each of the cavities 130 has a shape that is complementary to a shape of the analyte 10, like a lock-key interrelationship. The cavity internal wall 131 is made from a material that also contains the non-metallic element. In preferred embodiments, at least some of the non-metallic elements in the cavity internal wall material are directly exposed (i.e. not buried inside the material) to the cavity space, to facilitate the interaction or affinity between the non-metallic elements in the cavity internal wall material and the non-metallic elements in the analyte within the cavity.

With reference to FIG. 2A, various embodiments of the invention provide a sensing device 001 that further includes a reference sensor 200. Sensor 200 is the same as the working sensor 100 except that (1) the reference sensor 200 does not include the cavities 130 as those in the working sensor 100, and (2) voids 220's size of the reference sensor 200 is different from (bigger than or smaller than) voids 120's size of the working sensor 100.

In various exemplary embodiments, the non-metallic element may be selected from F, Cl, Br, I, O, S, Se, Te, N, P, As, Sb, B, C, H, or any combination thereof, among which F, Cl, Br, I, O, S, Se, Te, N, P, As, and Sb are preferred due to strong intermolecular interaction or affinity between electronegative elements e.g. F—F, Cl—Cl, Br—Br, I-I, O—O, S—S, Se—Se, Te—Te, N—N, P-P, As—As, and Sb—Sb.

In various exemplary embodiments, the sensing body, the void internal walls, and the cavity internal walls may be made from same or different material. In preferred embodiments, the sensing body, the void internal walls, and the cavity internal walls are all made from a same material containing the non-metallic element. For example, such same material may comprise a polymer prepared from photo polymerization and/or thermal polymerization using monomers containing the non-metallic element. In a specific embodiment, such same material is prepared from a pre-polymerization composition comprising the monomers containing the non-metallic element, the analyte containing the non-metallic element, and an optional cross-linking agent. For example, the pre-polymerization composition may include template/analyte molecule PFOA; functional monomers including 2-(trifluoromethyl) acrylic acid (TFMAA), 2-(difluoromethyl) acrylic acid (DFMAA), and/or 2-(monofluoromethyl) acrylic acid (MFMAA); and cross-linking agent EGDMA that utilizes an interaction between the non-metallic elements such as fluorine-fluorine interactions, electrostatic attraction, and associated weak interactions. In some embodiments, the pre-polymerization composition further comprises monomers that do not contain the non-metallic element such as acrylic acid (AA), methyl acrylic acid (MAA), and any mixture thereof.

In various exemplary embodiments, the array of voids is a 3D array of voids formed by removing a colloidal crystal from a solid body into which the colloidal crystal is incorporated and integrated. For example, the colloidal crystal may include silica nanoparticles, polystyrene nanoparticles, polymer beads, oxides, oxide ceramic nanoparticles, nitrides, carbides, quantum dots, macromolecules, carbon nanostructures, or any combination thereof. The particles assembled in one of the monolayers (2D arrays) in the colloidal crystal may be the same as, or different from, the particles assembled in another one of the monolayers (2D arrays). The size (diameter) of the voids may be in the range of 10-1000 nm, 50-1000 nm, 50-500 nm, 150-300 nm, or 180-400 nm.

In various exemplary embodiments, the 3D array of voids may be formed by stacking a number of 2D array of voids, and a height of the stack of 2D array of voids, or a thickness of the 3D array of voids, may be approximately 2-10 μm. For example, the 3D array of voids may be formed by stacking 5-20 (e.g. 10) layers of 2D array of voids. The 2D array of voids (e.g. measured from the top layer) may have a uniform area of 0.01-4 cm² such as larger than 2×2 mm² and up to 2×2 cm².

In various exemplary embodiments, the sensing body is deposited on the base 73 as shown in FIG. 1I, for example, base 71 may be a polymer plate such as a PMMA plate.

In various exemplary embodiments, a binding of the analytes to the cavities induces or triggers a detectable variation of the optical property of the 3D array of voids, including the spectrum of light that is transmitted through, reflected from, and/or diffracted from the 3D array of voids; and a degree of the detectable variation is correlated with the amount of the analytes bound to the cavities. The sensing device of the invention may include a light source such as a laser emitting light e.g. a light beam that irradiate upon the 3D array of voids (as incident light). A spectrometer may be then used to measure the spectrum of light that is transmitted through, reflected from, and/or diffracted from the 3D array of voids. A computer may be used to record and analyze the obtained spectrum or spectra.

In various exemplary embodiments, the present invention provides a method of measuring an analyte containing a non-metallic element using the sensing device as described above. As shown in FIG. 2B, the method includes (i) contacting a sample of the analyte with the working sensor, (ii) binding the analyte to the cavities and inducing or triggering a detectable variation of the optical property of the 3D array of voids, including a spectrum of light that is transmitted through, reflected from, and/or diffracted from the 3D array of voids, and (iii) correlating a degree of the detectable variation to an amount of the analytes bound to the cavities.

For example, the analyte may contain F, C, and/or H. In specific embodiments, the analyte is selected from fluorinated chemicals such as perfluorinated chemicals (PFCs), e.g. perfluoroalkyl substance, for example, perfluorooctane sulphonate (PFOS) and perfluorooctanoic acid (PFOA); an herbicide such as atrazine, and PFAS (EPA 537).

In representative and still exemplary embodiments, the present invention provides a platform sensing technology for field trace detection of analytes 10 such as PFOA and PFOS in groundwater. A representative embodiment of the working sensor 100 is a photonic crystal-based polymer sensing chip with cavities 130 such as molecularly imprinted (MIP) binding sites, which can selectively bind to PFOA and PFOS molecules in water and produce specific optical signals that are read using a portable UV-Vis spectrometer and correlated to the concentration of PFOA and PFOS in water. The “molecularly-imprinted photonic crystal” can show the colors of rainbow through nano and molecular engineering. Such a working sensor 100 exhibits the following advanced attributes: high sensitivity, high specificity, fast detection, ease of operation, low power consumption, zero chemical release, and low operation cost. Moreover, direct use of the sample water eliminates any uncertainty associated with measurement technique or complicated separation processes. For example, such a MIP sensor 100 may be used for detecting herbicide atrazine in water.

As a representative and still exemplary embodiment of the 3D array of voids 120, a photonic crystal sensing chip 31 consists of a 3D-ordered interconnected macroporous structure. For ease of use, the chip 31 may be assembled onto a clear microscope slide support 32, as shown in FIG. 3. In the photonic crystal structure, numerous nanocavities (as an embodiment of cavities 130 in FIG. 1J) derived from removed analytes 10 such as template molecules (PFOA/PFOS in this embodiment) may be distributed in the thin walls (as an embodiment of cavity internal wall 131 in FIG. 1J) of the ordered macro-pores (inverse polymer opal). During detection, these nanocavities will recognize the template molecules with high specificity and induce a change in the refractive index of the ordered structure. The color of the sensor will change via Bragg diffraction, which can be detected using a portable UV-Vis spectrometer (a reader). The color change (i.e. absorption peak shift) of the sensor will be correlated with the concentration of template molecule in water. Based on this embodiment, an autonomous PFOA and PFOS monitoring system can be developed, including water sampling, measurement, data processing, and reporting.

With no intention of being bound by any particular theory, it is believed that once molecular recognition occurs, the trapped analyte molecules (i.e. binding of analyte 10 to cavity 130) will cause either swelling or shrinkage of the prepared hydrogel, leading to refractive index change. The refractive index change of the sensing element will induce its diffraction peak shift, which can be detected optically and correlated with the concentration change of PFOA in water. The diffraction peak, λ_(max), for the porous hydrogel is determined by the Bragg equation (1):

$\begin{matrix} {\lambda_{\max} = {1.633\left( \frac{d}{m} \right)\left( \frac{D}{D_{0}} \right)\left( {n_{a}^{2} - {\sin \; \theta^{2}}} \right)^{1/2}}} & (1) \end{matrix}$

where d is the sphere diameter of the silica colloidal particle (which is also one way to define voids 120's size of the working sensor 100), m is the order of Bragg diffraction, (D/D₀) is the degree of swelling of the gel (D and D₀ denote the diameters of the gel in the equilibrium state at a certain condition and in the reference state, respectively), n_(a) is the average refractive index of the porous gel at a certain condition, and θ is the angle of incidence. According to this equation, if the molecular recognition process could cause swelling or shrinkage of the prepared hydrogel, then the readable optical signal is detectable.

In a representative and still exemplary embodiment, the fabrication procedure for a hierarchical porous sensor chip sensing chip is schematically shown in FIG. 4. The procedure starts with the preparation of a colloidal crystal template or array, followed by the infiltration and polymerization of the pre-ordered complex of PFOA with functional monomers (pre-polymerization complex) in the inter-spacers of the colloidal crystal, and then the removal of the used templates (colloid particles and PFOA/PFOS molecules). The feasibility of MIP photonic crystal-based sensing has been demonstrated by Zhen Wu et al, “Label-free colorimetric detection of trace atrazine in aqueous solution by using molecularly imprinted photonic polymers” Chemistry—A European Journal, v 14, n 36, p 11358-11368, which is incorporated herein by reference. In their approach, colloidal crystal templates used to form inverse polymer opals were created using a solvent evaporation colloidal crystal growth method, which normally requires several days. In contrast, the present invention provides a scalable two-phase assembly and transfer technique to fabricate colloidal crystal templates, which significantly reduces the preparation time to less than one hour.

Referring to FIG. 4, the procedure for sensing chip fabrication includes the following steps. (1) Preparation of colloidal crystal arrays: Silica colloidal crystal arrays are prepared using a two-phase self-assembly and transfer process to form highly ordered 3D macroporous structures. The size of monodispersed silica particles may range 10-1000 nm, 50-1000 nm, 50-500 nm, 150-300 nm, or 180-400 nm. Self-assembled monolayers of silica particles are stacked onto a glass or oxide ceramic support to form a multi-layer photonic crystal film with a thickness of approximately 3-5 μm. (2) Infiltration and polymerization of pre-polymerization complex: In order to fabricate the PFOA-imprinted polymer hydrogel, the template molecule (PFOA), functional monomers (TFMAA), and cross-linking agent (EGDMA) are first mixed to generate a pre-polymerization cluster that utilizes fluorine-fluorine interactions, electrostatic attraction, and associated weak interactions. The mixture is then be filled into the void spaces of the colloidal crystal array via capillary force by using a sandwich structure of PMMA/nanoparticle array/silica. Upon polymerization, the structure is frozen in a 3D network of polymers. (3) Removal of template particles and molecules: The removal of silica particles and the embedded PFOA molecules from the imprinted polymer matrix yields highly-ordered 3D and interconnected macroporous arrays with specific nanocavities that interact with PFOA molecules through non-covalent interactions.

In preferred embodiments, the invention provides (1) a two-phase assembly method to fabricate colloidal crystal templates with a uniform area larger than 1 cm²; (2) Preparation of inverse polymer opal sensors with specific binding sites for PFOA molecules using a fluorous monomer and cross-linker; (3) Fabrication of molecularly imprinted photonic crystal sensors on PMMA support with a uniform area larger than 1 cm²; and (4) a demonstration of a reproducible calibration curve for a range of concentrations from 0 or 0.1 to 1000 ppt of PFOA in water. Specifically, silica nanoparticle colloidal crystal templates have been fabricated using a two-phase interface assembly method. A fabrication process for the molecularly imprinted sensor has been developed. A detection limit of 10 ppt has been achieved for trace detection of PFOA in a mixed solvent of water/methanol in a lab setting.

In specific embodiments, the colloid crystal template was prepared using water-hexane interfacial assembly. As shown in FIG. 5, multi-layer colloid crystal templates were prepared on glass or oxide ceramic supports using a water/hexane interfacial assembly and transfer method. The process was started by positioning a pre-cleaned silica substrate on the bottom of a Petri dish. Then a suitable amount of water was poured into the Petri dish to immerse the substrate, followed by adding a few drops of hexane onto the water surface to form a thin organic solvent layer. A monolayer of silica colloidal nanoparticles was prepared by spreading the ethanol suspension (ca. 1% w/w) of silica nanoparticles onto the interface between the water and the thin layer of hexane in the Petri dish until the water surface was totally covered with a bead monolayer. Then the monolayer was lowered onto the substrate by decreasing the water level and allowing the water to flow out of the dish through a valve on the bottom. Multi-layer colloid crystal templates were fabricated by repeating this process until the desired number of layers was reached. Two kinds of silica nanoparticles with particle sizes of 180 nm and 300 nm were used to make colloid crystal templates with up to 10 layers. Two different glass substrates were used as support, including a circular disc of 18 mm in diameter and 0.17 mm in thickness and a square of 18 mm×18 mm with a thickness of 0.17 mm. Steps (a) and (b) in FIG. 5 are spreading beads from ethanol dispersion on the water-hexane interface; step (c) is the formation of the bead monolayer; and step (d) is transfer of the bead monolayer onto glass or oxide ceramic substrate.

With reference to FIG. 6, silica nanoparticle colloids with different particle size in panel a) were purchased from NanoComposix (San Diego, Calif.) in colloid form. NanoComposix's silica nanospheres are monodisperse with diameters 20 nm and up, and are available with both bare and amine-terminated surfaces. Panels b) and c) show that ethanol-dispersed monodisperse silica nanoparticles of 180 nm and 300 nm were used. Panel b) shows a TEM image of the 180 nm silica particles, and Panel c) shows size distribution of particle obtained from multiple TEM images.

Multi-layer colloid crystal templates of 180 nm silica nanoparticles were fabricated by repeating the monolayer deposition process. The samples were annealed at 70° C. for 10 minutes after the deposition process to evaporate trapped water, hexane or ethanol from the template. It was found that this annealing process increases the adhesion between the particles and the glass or oxide ceramic substrate. FIG. 7 shows SEM surface morphologies of a 1-layer sample in panel (a) and 10-layer sample in panel (b) under the same magnification. It can be seen from this figure that a certain degree of order is retained in the 10-layer sample with a sufficient domain size.

The UV-Vis absorption spectra of a 5-layer and a 10-layer colloid crystal template were measured using a UV-Vis spectrometer. FIG. 8 shows the UV-Vis absorption spectra of a 5-layer and a 10-layer silica colloid crystal template with a particle size of 180 nm. As can be seen from the figure, there is no obvious absorption peak on the spectrum of the 5-layer sample (red curve). For the 10-layer sample, an absorption peak can be clearly seen on the spectrum (green curve). The absorption peak is believed to be caused by optical interference between the ordered nanoparticle layers. Optical interference is stronger in the 10-layer sample than in the 5-layer sample, which is why the absorption peak is more obvious in the 10-layer sample.

Colloid crystal templates were also fabricated using 300 nm silica nanoparticles using a procedure similar to the one described above for 180 nm silica nanoparticles. FIG. 9 Panel (a) shows a 10-layer colloid crystal sample of 300 nm silica nanoparticles on 18 mm×18 mm glass substrate, and FIG. 9 Panel (b) shows vis-NIR spectra of the colloid crystal sample measured after assembly of 5 layers and 10 layers. A typical 10-layer colloid crystal sample is shown in FIG. 9 Panel (a) with vis-NIR spectra of this sample measured after assembly of 5 layers and 10 layers shown in FIG. 9 Panel (b). As can be seen from FIG. 9 Panel (b), the 300 nm sample possesses a strong absorption peak at 679 nm, which may be due to strong reflection from the ordered layers. This reflection in the red region of the solar spectrum results in the reddish appearance of the sample under white illumination (as shown in FIG. 9 Panel (a)). Comparison of spectra of the 180 nm and 300 nm samples indicates that increasing particle size from 180 nm to 300 nm causes a red shift of absorption peak from 438 nm to 668 nm.

The fabrication process for the PFOA-imprinted inverse polymer opal structure was provided in an exemplary embodiment. PFOA-imprinted photonic polymer hydrogels were fabricated over the colloid crystal templates as shown in FIG. 10. Monomer solutions were poured over the opal structured 3D ordered array in order to obtain inverse opal replicas of a self-assembled colloidal crystal template via a noncovalent, self-assembly approach. The template molecule (PFOA), monomer (TFMAA), and cross-linking agent (EGDMA) were first mixed to generate a pre-polymerization cluster that utilizes fluorine-fluorine interactions, electrostatic attraction, and associated weak interactions.

In a typical sensor preparation, PFOA, TFMAA and EGDMA were mixed in methanol at a molar ratio of 1:2:2 and left overnight to allow sufficient complexation. Due to hydrogen bonding and fluorine-fluorine interaction, good dispersion of the analyte molecules in the matrix was achieved. Then, 3 wt % of AIBN was added as a radical initiator to initiate free radical polymerizations and the mixture was degassed with nitrogen for 10 min. A suitable amount of the well-dispersed monomer solution was poured over a colloid crystal template, which was then covered with a PMMA plate to form a sandwich structure. Once the colloidal crystal layer in the formed sandwich structure became transparent, a successful infiltration process was completed. After the removal of excess precursors, photo-polymerization was carried out under UV light at 365 nm for 2 h. The sandwich structure was then immersed in 20 wt % hydrofluoric acid for 2 hrs to fully etch the glass substrate and silica nanoparticles. The formed inverse opal polymer layer remained on the PMMA plate. The embedded PFOA molecules were removed by incubating the polymer film in an acetic acid/methanol mixture for 30 min, followed by drying in an ambient environment. A fabricated inverse opal sensor from a colloid crystal template of 300 nm nanoparticles is shown in FIG. 1I. For control experiments, non-imprinted photonic hydrogel (NIPP) films may be prepared by using the same procedure and conditions, only without the addition of PFOA molecules. FIG. 1I shows a fabricated inverse opal sensor from a colloid crystal template of 300 nm nanoparticles.

UV cross-linking of the mixed precursors was investigated using FTIR spectroscopy. After 2 hours of UV polymerization, the sample was measured using a FTIR to identify spectral peaks of molecular groups in the polymer sample. FTIR spectra of a UV-cured polymer (red) and the mixed precursor solution (black) are shown in FIG. 12. From these spectra, it can be clearly seen that the vibration peaks at 1637 cm⁻¹ of the C═C groups were significantly reduced in the polymer sample compared with that in the spectrum from the precursor solution. This means that the UV curing conditions can polymerize the monomer solution into a polymer. FIG. 12 shows the FTIR spectra of a UV-cured polymer (red) and the mixed precursor solution (black), with C═C vibration peak showing at 1637 cm⁻¹.

The surface morphologies of the fabricated reverse opal polymer structures were observed using SEM. FIG. 13 shows the surface morphologies at different magnifications. As can be seen from these images, the glass or oxide ceramic support and silica nanoparticles have been completely etched away. FIG. 13 includes surface morphologies of an inverse opal polymer structure at different magnifications (panels (a) and (b)) and at the edge of the structure (panel (c)).

The vis-NIR spectra of the fabricated inverse opal sensor after HF etching (red) and after PFOA removal (green) are shown in FIG. 14, together with a vis-NIR spectrum of the template for comparison (blue). After UV polymerization of the mixed precursors, the fluoro-containing polymer has a lower refractive index, leading to blue shift of the absorption peak (from 679 nm to 604 nm) for the etched opal structure compared with that of the template. After PFOA removal, there is further blue-shifting of the absorption peak to 569 nm, which is caused by the refractive index reduction from PFOA removal. FIG. 14 shows Vis-NIR spectra of the inverse opal sensor after HF etching (red) and after PFOA removal (green), a vis-NIR spectrum of the template is given for comparison (blue).

Trace detection of PFOA was successfully demonstrated using the PFOA-imprinted photonic crystal sensor. Due to the low surface energy of the fluoro-containing sensor, a suitable amount of methanol was added to the PFOA water solution for improved analyte/sensor affinity. During measurement, 5-10 μL of PFOA solution was pipetted over a 10×10 mm sensing area. After incubating for 10 seconds, the solution on the sensor was wiped away. After the solvent evaporated, the vis-NIR spectrum of the sensor was measured and the absorption peaks and corresponding wavelengths were identified. PFOA solutions with PFOA concentrations of 10 ppt, 20 ppt, 100 ppt, 1000 ppt and 10 ppb were prepared using a mixed solvent of water/methanol (1/4, v/v). Extensive sensor evaluation, including other test solutions with reduced methanol content (or other environment-friendly solvents such as ethanol and IPA) may be performed as well. The sensors were fabricated using colloid crystal templates of 300 nm silica nanoparticles. FIG. 15A shows vis-NIR spectra of the sensor at different concentrations from 10 ppt to 10,000 ppt. FIG. 15A shows Vis-NIR spectra of the molecularly imprinted sensor at different PFOA concentrations from 10 ppt to 10,000 ppt. It can be clearly seen that the absorption peak red-shifts with increasing PFOA concentration, as indicated in Table 1 below. Table 1 lists the absorption peak wavelengths at different PFOA concentrations. FIG. 15B shows a calibration curve in the range from 0 ppt to 10,000 ppt.

TABLE 1 PFOA Concentration Peak Wavelength (ppt) (nm) 0 553 10 564 20 566 100 568 1,000 571 10,000 573

For comparison, non-molecularly imprinted sensors were fabricated and evaluated. The vis-NIR spectra of the sensor at different PFOA concentrations from 10 ppt to 1,000 ppt are given in FIG. 16. The absorption peak wavelengths at different PFOA concentrations are listed in Table 2. As expected, no absorption peak shift was observed for the non-molecularly imprinted sensor in the detected concentration range of PFOA.

TABLE 2 PFOA Concentration Peak Wavelength (ppt) (nm) 0 544 10 544 20 543 100 543 1000 544

Various embodiments of the invention can provide a facile technology for fabricating a highly-ordered 3D colloidal photonic crystal array that consistently produces very good optical signals. Multiple highly-ordered multi-layered colloid crystal templates were prepared on a glass or oxide ceramic slide using this two-phase self-assembly process. Maximum thickness of the template was approximately 3 μm, with a uniform area larger than 3 cm². This crystal template was used to create a 3D-ordered and interconnected macroporous MIP structure which was utilized to detect PFOA molecules inside a solution. Polymer photonic crystal sensors successfully detected the 10 ppt target molecule solutions of PFOA by showing excellent Bragg peak shift (over 10 nm). MIP-based sensors could function similarly to pH paper and provide a promising alternative for rapid monitoring of PFOA levels on the spot. The MIP sensor has sufficient specificity, and it can be used in arrays of optical sensors in which each individual sensor can detect a different PFAS.

Referring back to FIG. 2, various embodiments of the invention provide a sensing device 300 comprising a working sensor 100 as described above. In preferred embodiments, sensing device 300 further includes a reference sensor 200 that is the same as the working sensor 100 except that (1) the reference sensor 200 does not include the cavities 130 as those in the working sensor 100, and (2) voids 220's size of the reference sensor 200 is different from (bigger than or smaller than) voids 120's size of the working sensor 100.

In representative and still exemplary embodiments, the present invention provides a bi-layered molecularly imprinted photonic crystal sensor with built-in standard. For example, a bi-layered MIP photonic crystal-based sensing device can achieve the goal of fast field trace detection of atrazine in water. The photonic crystal sensing chip includes two layers of 3D-ordered interconnected macroporous structure. The top layer (MIP layer) may be molecularly imprinted and have a longer periodical length (i.e. a longer absorption peak wavelength) while the bottom layer will be non-imprinted (non-MIP) and have a shorter periodical length (i.e. a shorter absorption peak wavelength). For ease of use, the chip may be assembled onto a clear microscope slide support, as shown in FIG. 17. In the MIP layer, numerous nanocavities derived from atrazine molecular imprinting (MIP) may be distributed in the thin walls of the ordered macro-pores (inverse polymer opal). During detection, the nanocavities will recognize atrazine molecules with high specificity and induce a refractive index change of the ordered structure, leading to an absorption peak wavelength shift. The peak shift can be detected using a handheld UV-Vis spectrometer, which can be correlated with the concentration of atrazine in water. The non-MIP bottom layer cannot bind to the analyte molecules, so its absorption peak wavelength will not be affected by changing of the analyte concentration. The absorption peak from the non-MIP layer can be used as an internal standard to calibrate the peak shift of the sensing MIP layer. The embodiment thus provides a bi-layered MIP sensing platform with built-in standard (reference peak) to eliminate non-molecular binding induced effects.

During measurement, the sensing chip may be brought into contact with the analyte solution. The analyte molecules will bind to the MIP nanocavities, leading to absorption peak shift. Besides the peak shift induced by the binding of the analyte molecule other factors might also affect peak shift. Saturation of the polymer matrix by the solvent might induce swelling to some extent. Additionally, variations in the surrounding temperature may cause hydrogel contraction/expansion during the measurement. These non-molecular binding factors may also affect the absorption peak shift in the non-MIP layer. With this built-in standard, the peak shift effects from sources besides molecular binding can be eliminated. FIG. 18 shows the UV-Vis spectra of colloid crystals independently (Green and Red) and when combined (yellow). The sensor measures absorption peak wavelength shift instead of peak intensity change. Besides Bragg diffraction, peak intensity is also affected by other experimental factors such as substrate scattering, surface roughness, spectrometer settings, ambient light, etc. However, the absorption peak position is only determined by the periodical length of the inverse opal structure and the refractive index change of the hydrogel induced by the nanocavities binding template molecules. The other experimental factors mentioned above (which change peak intensity) won't change peak position.

There is no labeling process involved with the test process. Conventional immunoassays use labeled antibodies for the detection of biomolecules, which are time-consuming to create and expensive. Label-free detectors for biomolecules have drawn increasing interest from researchers in the fields of proteomics, clinical diagnostics, and environmental monitoring. In label-free sensors, target molecules are detected in their natural forms without any labeling process. The sensor of the invention has the advantage of eliminating time-consuming and expensive labeling steps, as well as allowing for kinetic measurement of molecular interactions. As artificial antibodies, molecularly imprinted polymers have high affinity for their template molecules. A limit of detection as low as 10⁻¹⁶ M of atrazine in water (equivalent to 0.0215 ppt) has been demonstrated. The high degree of specificity stems from the high affinity of molecularly imprinted nano-cavities.

Molecularly imprinted photonic polymers may be fabricated using the above procedure with some modifications. In order to make a bi-layered sensor, a non-MIP reference layer will first be made on a PMMA support, followed by a MIP sensing layer on the reference layer.

Using a bi-layer MIP sensor, the UV-vis spectrum of an atrazine solution may show two absorption peaks with a reference peak at λ_(r) and a sensing peak at λ_(s). Before measurement, the sensor can be calibrated with the solvent used to make the atrazine solution as the starting point, which gives a reference peak position at λ_(r0) and a sensing peak position at λ_(s0). During measurement of test solution #1 with atrazine concentration of C₁, its UV-Vis spectrum will show a reference peak position at λ_(r1) and a sensing peak position at λ_(s1). The absorption peak shift Δλ₁ can be calculated using the following equation:

Δλ₁=(λ_(s1)−λ_(r1))−(λ_(s0)−λ_(r0))  (2)

For test solution #n with atrazine concentration of C_(n), its UV-Vis spectrum will show a reference peak position at λ_(m) and a sensing peak position at λ_(sn). The absorption peak shift Δλ_(n) can be expressed as follows:

Δλ_(n)=(λ_(sn)−λ_(rn))−(λ_(s0)−λ_(r0))  (3)

With the known concentrations (C_(n)) of test solutions and their corresponding absorption peak shift (Δλ_(n)), a calibration curve can be plotted. From the calibration curve, the limit of detection, sensitivity, and dynamic range of the sensor can be deduced. In order to determine the atrazine concentration of an unknown solution, the solution will be applied on a sensor with a known calibration curve. The UV-Vis spectrum of the sensor will be measured and its atrazine concentration can be determined from the absorption peak shift.

In other specific embodiments, the present invention provides a molecular imprinted photonic crystal sensor, and a novel fabrication process thereof, which includes a step to fabricate a multi-layered silica nanoparticle template. The embodiments can achieve the goal of fast field trace detection of e.g. atrazine in water. The developed photonic crystal sensing chip consists of a 3D-ordered interconnected macroporous structure. For ease of use, the chip is assembled onto a clear microscope slide support 32, as shown in FIG. 3. In the photonic crystal structure, numerous nanocavities derived from atrazine molecular imprinting (MIP) are distributed in the thin walls of the ordered macro-pores (inverse polymer opal). During detection, the nanocavities can recognize atrazine molecules with high specificity and induce a refractive index change of the ordered structure. The color of the sensor changes via Bragg diffraction, which can be detected using a handheld UV-Vis spectrometer (a reader). The color change (i.e. absorption peak shift) of the sensor is correlated with the concentration of atrazine in water. The embodiments provide an easy-to-use “Cide Counting” evaluation kit, including a sensing chip, a chip reader, a cartridge, and accessories. Once molecular recognition occurs, the trapped analyte molecules will cause either swelling or shrinkage of the prepared hydrogel, leading to refractive index change. The refractive index change of the sensing element will induce its diffraction peak shift, which can be detected optically and correlated with the concentration change of atrazine in water. The diffraction peak, λ_(max), for the porous hydrogel is determined by the Bragg equation (1) as described above.

A specific method of the invention for fabricating a template using oil-water interface assembly may include the steps of: forming an oil-water interface; positioning a substrate underneath the interface; formulating ethanolic nanoparticle colloid; dipping a small droplet of the colloid at the interface; accomplishing monolayer arrangement of silica nanoparticles at the interface; lowering the monolayer on the substrate; thermally treating the substrate at an elevated temperature; and repeating the process for deposition of the next layer. In the method, the volume of the droplet is calculated using the area of the interface, the concentration and particle size of the colloid for monolayer coverage. The elevated temperature may be from room temperature to 200° C., from 50 to 150° C., or from 70 to 100° C.

Another specific method of the invention for fabricating a molecular imprinting (MIP) sensor may include the steps of: formulating of a mixture solution comprising monomer, cross-linker, initiator, template molecules, and solvent; filling the gaps in the template using the mixture solution; positioning a plastic plate over the liquid filled template; squeezing out excess liquid with the plastic plate; cross-linking the monomer liquid under a UV radiation; separating the glass support from the plastic plate; etching the glass particles in the cross-linked monomer matrix using a HF solution. The mixture solution comprises template molecule, monomer, and cross-linker at a molar ration of, but not limited to, 1:4:1. The monomer can be, but not limited to, organic carbon or fluoro-carbon compounds with one of multi-—C═C bonds including acrylic acid (AA), methyl acrylic acid (MAA), and 2-(trifluoromethyl) acrylic acid (TFMAA). The template can be, but not limited to, organic carbon, fluorine or fluoro-carbon compounds including atrazine and perfluorinated chemicals (PFCs). The plastic plate can be, but not limited to, PMMA. The UV radiation wavelength can be, but not limited to, 365 nm. The HF concentration can be, but not limited to, 1 to 10 wt %.

In a specific embodiment, the fabrication starts with the preparation of a colloidal-crystal template, followed by the infiltration and polymerization of the pre-ordered complex of atrazine with functional monomers in the inter-spacers of the colloidal crystal, and then the removal of the used templates (colloid particles and atrazine molecules). The embodiments provide an in-house developed scalable two phase assembly and transfer technique to fabricate colloidal crystal templates, which will significantly reduce preparation time to less than one hour.

Silica colloidal crystal arrays are prepared using an in-house developed two-phase self-assembly and transfer process to form highly order 3D macroporous structure. The size of monodispersed silica particles used in this embodiment may range from 150 nm to 200 nm. Self-assembled monolayers of silica particles are stacked onto a glass or oxide ceramic support to form a multi-layer photonic crystal film with a thickness of about 2 μm. Regarding infiltration and polymerization of pre-polymerization complex, in order to fabricate atrazine-imprinted polymer hydrogel, the template molecule (atrazine), functional monomer (methacrylic acid, MAA), and cross-linking agent (ethylene glycol dimethylacrylate, EGDMA) are first mixed to generate a pre-polymerization cluster that utilizes hydrogen-bond interactions, electrostatic attraction, and associated weak interactions. The molecular structures of (a) atrazine, (b) MAA, and (c) EGDMA are shown below. The mixture is then filled into the void spaces of the colloidal-crystal array via capillary force by using a sandwich structure of PMMA/nanoparticle array/silica (as shown in FIG. 4). Upon polymerization, the structure is frozen in a 3D network of polymers. The removal of silica particles and the embedded atrazine molecules from the imprinted polymer matrix yields highly ordered 3D and interconnected macroporous arrays with specific nanocavities that could interact with the atrazine molecule through noncovalent interactions.

In the specific embodiments, a two phase interfacial self-assembly technique is developed to prepare colloid crystal templates, potentially reducing the preparation time down to less than an hour per template. Compared with the solvent evaporation method (normally lasting for a few days), this represents a significant step forward toward commercialization of the photonic crystal sensing technology. A closely packed 2-dimensional (2D) colloid crystal layer is self-assembled at the interface of water and oil phases. Then a colloid crystal template is built by stacking the closely packed 2D layers together through a layer-by-layer transfer approach. After transferring onto a substrate, the substrate with the assembled bead layer is thermally treated to increase coating robustness for subsequent transfers. The process is repeated until a desired number of layers is reached.

In an example, silica (200 nm) colloids were purchased from NanoComposix (San Diego, Calif.). NanoComposix's silica nanospheres are monodisperse with diameters 20 nm and up, and are available with bare and amine-terminated surfaces. The company can provide silica nanoparticles either as a dried powder or in solution at 10 mg/mL. Standard silica particles are provided in water and amine-terminated silica particles are provided in ethanol. Water dispersed mono-dispersed silica nanoparticles of 200 nm with certified variations (CVs) of <12% was used in this example.

Referring back to FIG. 5, oil-water interfacial assembly of silica colloidal nanoparticles was prepared using a process that starts by positioning a pre-cleaned silica substrate on the bottom of a Petri dish. Then a suitable amount of water is poured into the Petri dish to immerse the substrate, followed by adding a few drops of hexane or heptane onto the water surface to form a thin organic solvent layer. A monolayer of silica colloidal nanoparticles is prepared by spreading the EtOH suspension (ca. 5% w/w) of silica nanoparticles onto the interface between the water and the thin layer of hexane or heptane in the Petri dish until the water surface is totally covered with a bead monolayer. Then the monolayer is lowered onto the substrate by decreasing the water level and allowing the water to flow out of the dish through a valve on the bottom. This “water level lowering” method offers advantages over a “picking-up with a substrate” method, such as less interruption of coating morphology and ease of scaling-up.

Then, multi-layered silica nanoparticle templates were built by stacking the closely packed 2D monolayers together. After a monolayer was transferred onto the substrate, the substrate with the assembled bead layer was thermally treated to increase coating robustness for subsequent coatings. The process repeated until a defined number of layers are reached.

Subsequently, molecularly imprinted photonic polymers were fabricated. To achieve multipoint interacting binding sites of high selectivity in the resulting photonic polymer, excess functional monomer was added. In the specific embodiments, the molar ratio between atrazine, MAA, and EDGMA was fixed at 1:4:1. In a typical preparation for MIP polymer, atrazine (1 mmol, 216 mg), MAA (4 mmol, 344 mg) and EGDMA (1 mmol, 0.2 mL) was mixed in methanol (0.5 mL), and left overnight to allow sufficient complexation. Then, AIBN (6.1 mmol, 0.01 g) was added as a radical initiator to initiate free radical polymerizations, and the mixture was degassed with nitrogen for 10 min. The mixture was cast on a glass slide with a colloidal-crystal film. Once the template became transparent, the voids among silica nanoparticles were completely filled, as shown in FIG. 10. Then a PMMA plate was positioned on the top of the template and squeezed out excess liquid. After the removal of excess precursors, photo-polymerization was carried out in an ice bath under a UV light at 365 nm for 2 hours. After separating the glass substrate from the polymer, the polymer was immersed in 1% hydrofluoric acid for 2 h to fully etch the used silica particles. The formed MIP polymer films remained on the PMMA substrate. The embedded atrazine molecules were then removed by incubating the polymer film in an acetic acid/methanol mixture for 6 h, followed by rinsing with phosphate buffer solution (pH 7.6).

In some improved embodiments of the invention, the interface assembly process has been optimized. The colloidal substrate size was scaled up from 18 mm×18 mm to 24 mm×24 mm, then to 63 mm×63 mm. In the process, the container used for substrate assembly was scaled-up, and the amount of solvent and colloidal particle solution used was calculated. Substrate drying time was longer. For a larger glass support, more water/solvent was left on the support. It took more time for the water/solvent to dry using the drying process as described above, which was drying the support inside an oven at 60° C. for 7-10 minutes. In the beginning of the process, it took more than 20 minutes for substrates to dry using the drying process. Moreover, the assembled colloidal substrates possessed less order of particle arrangement. After the assembly of a nanoparticle layer, the assembled substrate will be transported into an oven for drying. After the low boiling point solvent evaporated, the water/solvent interface disappeared. Then, the longer the water drying process was, the more nanoparticles lost the ordered arrangement formed during the interface assembly process.

The improved embodiments provide solutions to solve these issues. First, this on-site drying process employed an IR lamp, and eliminated the transportation of the assembled substrate from an assembly chamber to a drying oven, thus simplifying the assembly process. It also avoided any mechanical vibration from handling the wet substrate during substrate separation from the support and transport from the chamber to the oven, which could damage the formed order of particles in the film. Second, solvent with a boiling point close to that of water was utilized. The solvent was switched from hexane to heptane which has a boiling point close to that of water. In such a solvent/water system, the interface can remain longer during the drying process. Therefore, more particles can maintain the ordered arrangement. In summary, using an IR lamp with suitable power and heptane as the solvent, colloidal substrates with highly ordered nanoparticle arrangement were fabricated and characterized. The drying process can be as fast as 6-8 minutes. This scale-up process has laid a foundation for further scale-up of the substrate to a multi-inch square dimension.

In the improved embodiments of the invention, switching to injection using a syringe pump in step (iii) of FIGS. 1C and 1E was made due to the enhanced control over factors such as pipet position, rate of distribution, and movement of the pipet, and to minimize human error. Using the previous manual distribution method, the depth and position of the pipet were difficult to stabilize, causing the pipet tip to routinely breach the heptane-water interface and create layers with poor order. In the method, a syringe pump with a constant injection rate was used to push a predetermined volume of silica nanoparticle solution through a syringe with its tip held in a fixed position. This delivery system led to higher quality layers with more order and greater reproducibility. Using the experiment set-up with a syringe pump injection and an IR lamp, an assembled silica nanoparticle multi-layer sample can be fabricated with consistent quality throughout the entire dimension, as can be observed from the typical Vis-NIR spectra from the 4 corner areas, as illustrated in FIG. 19.

In the improved embodiments of the invention, densely packed monolayer was made through compressing the layer with a pair of barriers, as shown in FIG. 1G. The interface assembly method allows forming a monolayer by dispersing a calculated amount of particle solution over a certain area, which might not be very densely packed. For some applications where densely packed particles are needed, a compressing step can be added using a pair of barriers to form a densely packed layer.

As described above, the particles 64 in FIGS. 1D and 1F may comprise silica nanoparticles, polystyrene nanoparticles, polymer beads, oxides, oxide ceramic nanoparticles, nitrides, carbides, quantum dots, macromolecules, carbon nanostructures, or any mixture thereof. The preparation of the particles 64 may be a suspension of the particles in an organic solvent such as ethanol, for example nanoparticle colloid mono-dispersed in ethanol. In the improved embodiments of the invention, the interface assembly technology can also be used for other building blocks such as polymer beads, oxides, nitrides, carbides, quantum dots, macromolecules, and carbon nanostructures. An important step of this process is to formulate stable ethanol based colloidal solutions of these building blocks. With that, ordered multi-layers of a single building block, and ordered multi-layers of different building blocks can be prepared. In other words, the particles 64 in FIGS. 1D and 1F assembled in one of the monolayers (65, 66, 67 . . . ) may be the same as, or different from, the particles 64 assembled in another one of the monolayers (65, 66, 67 . . . ).

In the foregoing specification, embodiments of the present invention have been described with reference to numerous specific details that may vary from implementation to implementation. The specification and drawings are, accordingly, to be regarded in an illustrative rather than a restrictive sense. The sole and exclusive indicator of the scope of the invention, and what is intended by the applicant to be the scope of the invention, is the literal and equivalent scope of the set of claims that issue from this application, in the specific form in which such claims issue, including any subsequent correction. 

1. A process of preparing a 3D array of particles by stacking multiple 2D arrays of the particles, comprising: (a) providing a container with a substrate inside the container; (b) introducing a first liquid into the container to immerse the substrate, and optionally introducing a second liquid to float on top surface of the first liquid and form a two-phase system; (c) assembling the particles on top surface of the first liquid to form a monolayer (2D array) of the particles thereon, or optionally assembling the particles to form a monolayer at an interface between the first liquid and the second liquid; (d) moving the monolayer and/or the substrate to reduce a distance therebetween until the monolayer is deposited on the substrate; (e) subjecting the substrate with the monolayer deposited thereon to steps (a)-(d) to stack/deposit another monolayer on top of the monolayer previously deposited; (f) optionally repeating step (e) until a desired number of monolayers are stacked on the substrate.
 2. The process according to claim 1, wherein a flat surface of the substrate for depositing the monolayer(s) is not vertical to the top surface of the first liquid; for example, the angle between the two surfaces may range from −89° to 89°, from −70° to 70°, from −50° to 50°, from −35° to 35°, from −20° to 20°, from −10° to 10°, from −5° to 5° such as substantially 0°.
 3. The process according to claim 1, comprising: (i) providing a container with a substrate on a bottom of the container; (ii) filling a first liquid into the container until the substrate is immersed in the first liquid; (iii) adding a preparation of the particles into the container, and assembling the particles on top surface of the first liquid to form a monolayer (2D array) of the particles thereon; (iv) removing or discharging the first liquid from the container so that said monolayer of the particles falls down onto the substrate, and is deposited thereon; (v) refilling the first liquid into the container until the substrate and the particles previously deposited thereon is immersed in the first liquid; (vi) repeating steps (iii) and (iv) so that another monolayer of the particles is deposited on the substrate by stacking over an immediate monolayer that has previously deposited thereon; and (vii) optionally repeating steps (v) and (vi) until a desired number of monolayers of the particles are deposited on the substrate.
 4. The process according to claim 3, comprising (i) providing the container with the substrate on the bottom of the container; (ii) filling the first liquid into the container until the substrate is immersed in the first liquid; (ii.5) adding a second liquid into the container, so that the second liquid is floating on top of the first liquid, wherein an interface is formed between the two liquids; (iii) adding the preparation of the particles into the container, and assembling the particles between the first liquid and the second liquid (or at their interface) to form the monolayer (2D array) of the particles; (iv) removing or discharging both the first liquid and the second liquid from the container so that said monolayer of the particles falls down onto the substrate, and is deposited thereon; (v) refilling the first liquid into the container until the substrate and the particles previously deposited thereon is immersed in the first liquid; (vi) repeating steps (ii.5), (iii) and (iv) so that another monolayer of the particles is deposited on the substrate by stacking over the immediate monolayer that has previously deposited thereon; and (vii) optionally repeating steps (v) and (vi) until a desired number of monolayers of the particles are deposited or stacked on the substrate.
 5. The process according to claim 4, wherein step (iii) further comprises compressing the monolayer of the particles with a pair of barriers to reduce the area of the monolayer and to pack the particles in the monolayer more densely or more intimately.
 6. The process according to claim 4, wherein said adding the preparation of the particles into the container in step (iii) is accomplished with a pipette, a syringe pump, or any combination thereof.
 7. The process according to claim 4, further comprising step (viii) annealing the deposited particles at an elevated temperature such as 50-100° C. e.g. 70° C. for a period of time such as 5-20 minutes e.g. 10 minutes, to evaporate a residue of the first liquid, the second liquid, and a liquid in the preparation of the particles, optionally wherein the annealing is accomplished by oven drying, IR lamp heating, or any combination thereof.
 8. The process according to claim 4, wherein the particles comprise silica nanoparticles, polystyrene nanoparticles, polymer beads, oxide ceramic nanoparticles, oxides, nitrides, carbides, quantum dots, macromolecules, carbon nanostructures, or any mixture thereof; wherein some particles assembled in a monolayer may be the same as, or different from, other particles assembled in the same monolayers; and wherein the particles assembled in one of the monolayers may be the same as, or different from, the particles assembled in another one of the monolayers.
 9. The process according to claim 4, wherein the preparation of the particles comprises a suspension of the particles in an organic solvent such as ethanol, for example nanoparticle colloid mono-dispersed in ethanol.
 10. The process according to claim 4, wherein the substrate comprises a material made of glass or oxide ceramic; wherein the first liquid comprises a hydrophilic liquid such as water; and wherein the second liquid comprises a hydrophobic liquid or oil such as hydrocarbon liquid such as hexane, heptane, or any mixture thereof.
 11. The process according to claim 4, wherein the particles have an average size in the range of 10-1000 nm, 50-1000 nm, 50-500 nm, 150-300 nm, or 180-400 nm; and wherein each monolayer of the particles has a thickness that is substantially the same as the particles' average size.
 12. The process according to claim 4, wherein the stack of the monolayers comprises 1-00, 5-50, 5-20, or 10-20 (e.g. 10) monolayers of the particles, and the stack has a height of approximately 2-10 μm.
 13. A process for preparing a sensing body of a working sensor for a sensing device useful for detecting an analyte containing a non-metallic element, comprising (1) fabricating a 3D array of particles according to claim 1; (2) infiltrating the 3D array of particles with a mixture of a solidifiable material and the analyte, wherein the solidifiable material comprises said non-metallic element too; (3) solidifying the solidifiable material with the analyte; and (4) washing away the 3D array of particles and the analyte, forming a sensing body including a 3D array of voids each having a void internal wall; wherein at least a part of the voids are interconnected to each other and are configured to expose to said analyte in a future sample, and admit said analyte into said at least a part of the voids; wherein void internal walls of said at least a part of the voids have cavities each having a cavity internal wall; wherein each of the cavities has a shape that is complementary to a shape of the analyte; and wherein the cavity internal wall is made from a material containing said non-metallic element.
 14. The process according to claim 13, wherein the non-metallic element is selected from F, Cl, Br, I, O, S, Se, Te, N, P, As, Sb, B, C, H, or any combination thereof; and wherein the sensing body, the void internal walls, and the cavity internal walls are all made from a same material containing said non-metallic element.
 15. The process according to claim 14, wherein said same material comprises a polymer prepared from photo polymerization and/or thermal polymerization using monomers containing said non-metallic element.
 16. The process according to claim 15, wherein said same material is prepared from a pre-polymerization composition comprising said monomers containing said non-metallic element, the analyte containing said non-metallic element, and an optional cross-linking agent.
 17. The process according to claim 16, wherein the pre-polymerization composition comprises template/analyte molecule PFOA; functional monomers including 2-(trifluoromethyl) acrylic acid (TFMAA), 2-(difluoromethyl) acrylic acid (DFMAA), and/or 2-(monofluoromethyl) acrylic acid (MFMAA); and cross-linking agent EGDMA that utilizes an interaction between the non-metallic elements such as fluorine-fluorine interactions, electrostatic attraction, and associated weak interactions; and optionally wherein the pre-polymerization composition further comprises monomers that do not contain said non-metallic element such as acrylic acid (AA), methyl acrylic acid (MAA), and any mixture thereof.
 18. The process according to claim 13, wherein the analyte is selected from fluorinated chemicals such as perfluorinated chemicals (PFCs), e.g. perfluoroalkyl substance, for example, perfluorooctane sulphonate (PFOS) and perfluorooctanoic acid (PFOA); an herbicide such as atrazine, and PFAS (EPA 537).
 19. The process according to claim 13, wherein a binding of the analytes to the cavities induces or triggers a detectable variation of the optical property of the 3D array of voids, including the spectrum of light that is transmitted through, reflected from, and/or diffracted from the 3D array of voids; and a degree of the detectable variation is correlated with the amount of the analytes bound to the cavities.
 20. The process according to claim 13, further comprising preparing a reference sensor that is the same as the working sensor except that the reference sensor does not include the cavities as those in the working sensor, and the voids' size of the reference sensor is different from (bigger than or smaller than) that of the working sensor. 